Electroporation-Based Genetic Manipulation in Type I Methanotrophs

Abstract

Methane is becoming a major candidate for a prominent carbon feedstock in the future, and the bioconversion of methane into valuable products has drawn increasing attention. To facilitate the use of methanotrophic organisms as industrial strains and accelerate our ability to metabolically engineer methanotrophs, simple and rapid genetic tools are needed. Electroporation is one such enabling tool, but to date it has not been successful in a group of methanotrophs of interest for the production of chemicals and fuels, the gammaproteobacterial (type I) methanotrophs. In this study, we developed electroporation techniques with a high transformation efficiency for three different type I methanotrophs: Methylomicrobium buryatense 5GB1C, Methylomonas sp. strain LW13, and Methylobacter tundripaludum 21/22. We further developed this technique in M. buryatense, a haloalkaliphilic aerobic methanotroph that demonstrates robust growth with a high carbon conversion efficiency and is well suited for industrial use for the bioconversion of methane. On the basis of the high transformation efficiency of M. buryatense, gene knockouts or integration of a foreign fragment into the chromosome can be easily achieved by direct electroporation of PCR-generated deletion or integration constructs. Moreover, site-specific recombination (FLP-FRT [FLP recombination target] recombination) and sacB counterselection systems were employed to perform marker-free manipulation, and two new antibiotics, zeocin and hygromycin, were validated to be antibiotic markers in this strain. Together, these tools facilitate the rapid genetic manipulation of M. buryatense and other type I methanotrophs, promoting the ability to perform fundamental research and industrial process development with these strains.

INTRODUCTION

Methane, the principal component of natural gas and biogas, is a cheap, abundant, and renewable energy and carbon source. However, methane is also the second most prevalent greenhouse gas (1). Therefore, technologies to efficiently convert methane to chemicals or fuels can bring new sustainable solutions to a number of industries with large environmental footprints. Methane-oxidizing bacteria (methanotrophs) are able to use methane as their sole source of carbon and energy and thus are promising systems for methane-based bioconversion (2, 3). The bioconversion of methane to industrial products (single-cell proteins, biopolymers, lipids, etc.) using aerobic methanotrophs has been studied for approximately 50 years but has had little enduring success (4). Current biological engineering and systems biology approaches provide new opportunities for metabolic engineering of methanotrophs. However, to be an industrial workhorse like Escherichia coli, an industrial methanotroph needs to have a high growth rate, and efficient genetic tools for its manipulation and a fundamental knowledge base are needed.

Many methanotrophs have been isolated and characterized since the classic study of Whittenbury et al. (5). Aerobic methanotrophs are found in two major groups, the gammaproteobacterial methanotrophs (type I) and the alphaproteobacterial methanotrophs (type II) (2). Most studies and biotechnological efforts have focused on well-characterized species, such as Methylococcus capsulatus Bath, Methylosinus trichosporium OB3b, and Methylocystis parvus OBBP (6). In recent years, a variety of new strains have been isolated, including thermophilic, psychrophilic, acidophilic, alkaliphilic, and halophilic methanotrophs, which have expanded the physiological range of aerobic methanotrophs (7, 8). Moreover, some isolates show robust growth and promise for use in biotechnological applications. Type I methanotrophs are particularly well suited for use in industrial processes, as they utilize the highly efficient ribulose monophosphate pathway for formaldehyde assimilation into biomass (2, 6). A particularly promising type I methanotroph for industrial use is Methylomicrobium buryatense 5G, a haloalkaliphilic strain that was isolated directly on natural gas from a soda lake in the Transbaikal region of Russia, a harsh environment with fluctuating temperatures, salinities, and pH levels (7). The pH optimum of this strain is 9.5, and the optimum salt concentration is 0.75%. M. buryatense strain 5GB1, a lab-adapted variant of M. buryatense 5G, grows well in pure culture on natural gas, methane, and methanol (9). The genomes of M. buryatense 5G and 5GB1 were recently sequenced and annotated (10). All of these parameters make this strain well suited for development as an industrial system for the bioconversion of methane.

Historically, the slow pace of genetic manipulation has been one of the limiting factors in analyzing the gene functions of methanotrophic bacteria. At present, conjugation is a method commonly used for the transformation of methanotrophs (6). Recently, to metabolically engineer M. buryatense 5GB1, Puri et al. developed a number of genetic tools for use with the strain, including a sucrose counterselection system and a small replicable vector, and these were found to be sufficient for genetic manipulation (9). However, as these tools are based on conjugation and vector construction, a number of steps are necessary to carry out genetic manipulation, making current techniques time-consuming. To accelerate the use of modern metabolic engineering techniques in M. buryatense 5GB1 and other type I methanotrophs, more streamlined and versatile approaches are needed. Genetic manipulation via electroporation requires fewer steps than conjugation and creates the ability to transfer linear DNA fragments, which results in double-crossover deletion and insertion events and removes the need for the counterselection of E. coli postconjugation. In addition, it allows the direct transfer of genes that might be toxic in alternate hosts, such as E. coli, and avoids the alteration of plasmids occurring during conjugation (9). Electroporation protocols have been developed in Methylocystis sp. strain SC2 and Methylocella silvestris BL2, both of which are type II methanotrophs (11, 12). However, no electroporation method has been reported for type I methanotrophs, and previous attempts to demonstrate electroporation in M. buryatense were unsuccessful (9).

In this work, we report simple and rapid electroporation-based genetic manipulation tools for multiple type I methanotrophs, including electroporation of DNA fragments that were assembled by PCR. We were able to obtain a high efficiency of DNA transfer via electroporation in M. buryatense strain 5GB1C, Methylomonas sp. strain LW13, and Methylobacter tundripaludum strain 21/22, type I methanotrophs from three different genera with different physiological traits (9, 13), demonstrating the broad applicability of the techniques developed here. In M. buryatense, we also demonstrate marker removal after electroporation through sacB counterselection or FLP-FRT (FLP recombination target) site-specific recombination. These tools were validated by performing gene knockouts, long fragment deletion, and integration of a foreign fragment into the chromosome. These genetic tools will enable the rapid genetic manipulation and metabolic engineering of type I methanotrophs, accelerating the study of methanotrophs and their development for use in industrial processes.

MATERIALS AND METHODS

Bacterial strains, plasmids, oligonucleotides, and DNA manipulation techniques.

The bacterial strains and plasmids used in this study are listed in Table 1. All M. buryatense strains were derived from M. buryatense strain 5G (7). Strain 5GB1, a variant of M. buryatense 5G, shows a higher growth rate than its parent strain, and its sequence has several differences, identified by genome sequencing, from the published draft genome sequence of strain 5G (9). Strain 5GB1C, a variant of strain 5GB1 intentionally cured of its native plasmid, is capable of being conjugated with small IncP-based plasmids (9). The oligonucleotides (see Table S1 in the supplemental material) were synthesized by Integrated DNA Technologies (IDT; USA), and DNA sequencing was performed by Genewiz (USA). All plasmids were constructed by use of Gibson assembly (14). Phusion DNA high-fidelity polymerase was purchased from New England Biolabs (NEB; USA). A QIAquick PCR purification kit and a QIAquick gel extraction kit were supplied by Qiagen (Germany). Gene identities are from GenBank.

TABLE 1.

Strains and plasmids used in this study

Strain or plasmid	Characteristicsa	Reference or source
Strains
E. coli TOP10	F− mcrA Δ(mrr-hsdRMS-mcrBC) ϕ80dlacZΔM15 ΔlacX74 recA1 araD139 Δ(ara leu)7697 galU galK rpsL endA1 nupG	Invitrogen
E. coli BL21-AI	F− ompT hsdSB(rB− mB−) gal dcm araB::T7RNAP-tetA	Invitrogen
E. coli NEB C2925	ara-14 leuB6 fhuA31 lacY1 tsx-78 glnV44 galK2 galT22 mcrA dcm-6 hisG4 rfbD1 R(zgb-210::Tn10) Tets endA1 rspL136 dam-13::Tn9 xylA5 mtl-1 thi-1 mcrB1 hsdR2	NEB
Methylomonas sp. LW13	Type I methanotroph, wild type	13
Methylomonas sp. LW13-ΔglgA::km	Methylomonas sp. LW13 glgA::Kmr	This work
M. tundripaludum 21/22	Type I methanotroph, wild type	13
M. tundripaludum 21/22-ΔglgA2::km	M. tundripaludum strain 21/22 ΔglgA2::Kmr	This work
M. buryatense
5GB1	Variant of M. buryatense 5G	9
5GB1C	Variant of M. buryatense 5GB1 intentionally cured of its native plasmid and capable of being conjugated with small IncP-based plasmids	9
5GB1C-ΔglgA1::km	M. buryatense 5GB1C ΔglgA1::Kmr	This work
5GB1C-ΔglgA1	M. buryatense 5GB1C ΔglgA1	This work
5GB1C-ΔmmoD::km	M. buryatense 5GB1C ΔmmoD::Kmr	This work
5GB1C-ΔmmoD	M. buryatense 5GB1C ΔmmoD	This work
5GB1C-Δsmmo::km	M. buryatense 5GB1C ΔsMMO::Kmr	This work
5GB1C-Δsmmo	M. buryatense 5GB1C ΔsMMO	This work
5GB1C-Ppmo-xylE	M. buryatense 5GB1C with Ppmo-xylE inserted between the METBUDRAFT_2794 and METBUDRAFT_2795 genes	This work
Plasmids
pAWP89	An IncP-based broad-host-range plasmid containing dTomato driven by the tac promoter, Kmr	9
pCM433kanT	A broad-host-range sacB-based vector for unmarked allelic exchange, Kmr	9
pCM433-moD	pCM433kanT containing the deletion construct of mmoD	This work
pET-29b(+)	Expression vector, Kmr	Novagen
pET29-M	pET-29b(+) containing the METBUDRAFT_0046 gene, Kmr	This work
pFC25	A broad-host-range plasmid containing flp driven by the tac promoter, Hmr	This work
P7Z6	pMD18-T containing the zeocin resistance marker, Zeor	18
pCM158	A broad-host-range cloning vector, Kmr	20
pTEC27	Template for the hygromycin resistance marker, Hmr	Addgene (21)
pE-FLP	A temperature-sensitive plasmid containing the flp gene, Ampr	Addgene (22)
pCM130	xylE promoter-probe vector, Tetr	19

^aAmp^r, ampicillin resistance; Hm^r, hygromycin resistance; Km^r, kanamycin resistance; Tet^r, tetracycline resistance; Tet^s, tetracycline susceptibility; Zeo^r, zeocin resistance.

Culture, growth conditions, and phenotypic characterization.

All strains were cultured in an atmosphere of 25% methane in air at 30°C (M. buryatense 5GB1C) or 20°C (Methylomonas sp. LW13 and M. tundripaludum 21/22). The plates were incubated in sealed jars (Oxoid Limited, Hampshire, United Kingdom), while liquid cultures were grown in 250-ml glass serum bottles (Kimble Chase, Vineland, NJ, USA) sealed with rubber stoppers and aluminum seals (Wheaton, Millville, NJ, USA). Milli-Q H₂O (Barnstead) was used in this study. M. buryatense cultures were grown in NMS2 medium (9), and Methylomonas and Methylobacter cultures were grown in NMS medium (15). NMS medium and NMS2 medium were supplemented with 1.5% Bacto agar for plate growth. The following antibiotics were used for the selection of colonies of both methanotrophs and E. coli containing the correct genetic manipulations: kanamycin (Km), 50 μg/ml; rifamycin (Rm), 50 μg/ml; hygromycin (Hm), 100 μg/ml; and zeocin (Zeo), 30 μg/ml. To detect the activity of soluble methane monooxygenase (sMMO), M. buryatense strains were first grown in NMS2 medium without copper at 30°C for 20 h and then subjected to the colorimetric plate assay developed by Graham et al. (16).

Electroporation protocol for type I methanotrophs.

For cells grown on agar plates, one loopful of cells was spread across an entire NMS or NMS2 agar plate and grown under methane overnight (M. buryatense 5GB1C and Methylomonas sp. LW13) or for 2 days (M. tundripaludum 21/22). For cells grown in liquid, a 50-ml liquid culture was grown to stationary phase as described above. Cells were harvested from the liquid culture by centrifugation at 5,000 rpm and 4°C for 10 min. Cells were harvested from the plates by scooping the entire biomass from a plate. Cells were resuspended in 50 ml room temperature electroporation solution (H₂O, 9.3% sucrose, 10% glycerol, or 30% polyethylene glycol [PEG] 6000) and harvested by centrifugation at 5,000 rpm and 4°C for 10 min. The pellet was washed in 10 ml room temperature electroporation solution, transferred to a 15-ml conical tube, and centrifuged again at 5,000 rpm and 4°C for 10 min. The resulting pellet was resuspended in 100 to 150 μl electroporation solution and placed on ice. Fifty microliters of the cell suspension was mixed gently with DNA (usually less than 3 μl), and the mixture was transferred to an ice-cold 1-mm-gap cuvette (Bio-Rad). Electroporation was performed using a Gene Pulser II system (Bio-Rad) set at 25 μF and 200 Ω. Immediately following the electrical discharge, 1 ml of medium (room temperature) was added to the cells. The resuspended cells were then transferred into 10 ml medium in 250-ml serum bottles, which were crimp sealed, and then incubated with either 50 ml methane or 0.02% methanol. After incubation at 30°C (M. buryatense 5GB1C) or 20°C (Methylomonas sp. strain LW13 and M. tundripaludum 21/22) for 4 h or overnight with shaking, the cells were centrifuged at 5,000 rpm for 10 min at room temperature, resuspended in 1 ml medium, and spread onto selective plates.

Fusion of multiple DNA fragments by PCR.

Fusion of multiple DNA fragments by PCR was performed as described by Shevchuk et al. (17). In brief, it was carried out as follows. Overlaps of 30 to 40 nucleotides were introduced between each of two fragments through the use of primers. The fragments were amplified and gel purified. The reaction mixture in step A was 9.5 μl water, 5 μl Phusion buffer (5×) (NEB, USA), 2 μl deoxynucleoside triphosphate (dNTP) mix (2.5 mM each), 8 μl gel-purified fragments (about 50 ng each), and 0.5 μl Phusion DNA polymerase; the cycling parameters were an initial denaturation at 98°C for 2 min and subsequent steps of 98°C for 10 s, annealing at 55°C for 20 s, and extension at 72°C for 2 min for 16 cycles total. The reaction mixture in step B was 32 μl water, 10 μl Phusion buffer, 4 μl dNTP mix, 2 μl forward and reverse primers (10 mM) specific for the expected fragment, 1 μl of the unpurified PCR product from step A, and 1 μl Phusion DNA polymerase; the cycling parameters were an initial denaturation at 98°C 2 for min and subsequent steps of 98°C for 10 s, annealing at 60°C for 10 s, and extension at 72°C for 2 min for 32 cycles total. The resulting product was purified by use of a QIAquick PCR purification kit (Qiagen), and the purified product was directly electroporated into the strains.

Construction of ZS cassette.

The zeocin resistance marker (Zeo^r) was generated through two-step PCR amplification from plasmid p7Z6 (18) using primer pairs z51/z3 and z52/z3. The fragment containing the tac promoter (P_tac) and the Shine-Dalgarno (SD) sequence of lacZ from pCM66 (19) was introduced to the 5′ end of the zeocin resistance gene Streptoalloteichus hindustanus (Sh) ble with the primers z51 and z52. To assemble the Zeo^r-sacB (ZS) cassette, the sacB gene together with its putative SD sequence was fused to the 3′ end of Zeo^r with primers z52, z3, s5, and s3.

Introduction of Ppmo-xylE into the 5GB1C chromosome.

The pFC25 vector was constructed by use of Gibson assembly using a hygromycin-resistant version of the replicable vector pCM158 as a backbone (20). The hygromycin resistance marker (Hm^r) was amplified by PCR from plasmid pTEC27 (Addgene accession number 30182) (21), the flippase gene was amplified by PCR from plasmid pE-FLP (Addgene accession number 45978) (22), and P_tac was amplified from pAWP89 (9). To create the insertion PCR product containing the pmo promoter fused to xylE, two PCR products were generated for the flanking integration sites and correspond to the intergenic region and coding sequences of the genes METBUDRAFT_2794 and METBUDRAFT_2795. PCR products corresponding to the pmo promoter, xylE, and Zeo^r flanked by FRT sites were inserted between the flanking regions via the fusion PCR method. The xylE gene was amplified from pCM130 (19). FRT sites were appended to Zeo^r by amplifying Zeo^r with primers containing the FRT sites.

Deletion of R-M systems and in vitro methylation of plasmid DNA.

Ten restriction-modification (R-M) system-associated restriction endonuclease genes (see Table S2 in the supplemental material) were predicted to exist in the genome of M. buryatense 5G through analysis of the restriction enzyme database REBASE (23) and bioinformatics analysis. Among these restriction endonuclease genes, METBUDRAFT_0633, METBUDRAFT_3180, and METBUDRAFT_4274 (the gene locus tags in the Integrated Microbial Genomes system [http://img.jgi.doe.gov/]) were deleted through unmarked allelic exchange using plasmid pCM433kanT (9), and others were deleted by electroporating the fusion PCR product containing the kanamycin resistance marker and two flanking homology regions of the target gene.

The methyltransferase gene METBUDRAFT_0046 was ligated to pET-29b(+) between the NdeI and XhoI sites, generating pET29-M. This vector contains a His tag sequence that is added to the C terminus of the expressed protein. E. coli BL21-AI harboring pET29-M was grown at 37°C in Luria-Bertani broth supplemented with Km to an optical density at 600 nm of ∼0.5 and induced with 0.1% arabinose and 0.5 mM IPTG (isopropyl-β-d-thiogalactopyranoside) at 30°C for 4 h. His-tagged (C-terminal) protein was purified using Talon metal affinity resin (Clontech) according to the supplier's instructions. For in vitro methylation, 10 mg of purified protein was incubated with 50 mM Tris-HCl (pH 8.5), 50 mM NaCl, 80 mM S-adenosylmethionine, 10 mM dithiothreitol, 10 mM EDTA, and 10 μg plasmid DNA substrate in a 200-μl reaction mixture for 2 h, 4 h, or 8 h at 30°C. Methylated plasmid DNA was purified and concentrated by phenol-chloroform extraction and ethanol precipitation.

RESULTS AND DISCUSSION

Development of an efficient electroporation method for M. buryatense 5GB1C.

To facilitate the expansion of genetic tools in type I methanotrophs, work was initiated in M. buryatense 5GB1C. First, two new antibiotics, zeocin and hygromycin, were tested and found to be useful. In both cases, the 5GB1C strain was found to be sensitive to the antibiotics, and strains containing the resistance markers (Zeo^r and Hm^r) were found to be resistant at 30 and 100 μg/ml, respectively. Next, we developed an electroporation protocol using the 5GB1C strain. Shuttle plasmids extracted from E. coli TOP10 strains had been used in previous electroporation attempts, but no transformants were obtained under tested conditions (9). Restriction-modification (R-M) systems are a common barrier to DNA transformation in bacteria, and 10 R-M systems are predicted in the genome of M. buryatense 5G. Therefore, plasmids isolated from the 5GB1C strain were tested. Plasmid pAWP89 (Km^r) or pFC25 (Hm^r), isolated from strain 5GB1C, could be electroporated into strain 5GB1C at a frequency of approximately 5 × 10⁵ CFU per μg DNA. However, electroporation of the plasmids extracted from E. coli strains with (TOP10) or without (NEB C2925) Dam and Dcm methylation systems yielded no transformants. Furthermore, if plasmids harvested from strain 5GB1C were introduced back into E. coli, extracted, and electroporated into strain 5GB1C again, no transformants were obtained. Thus, these results support the hypothesis that R-M systems are a key limiting factor to the transformation of foreign DNA into strain 5GB1C.

Electroporation conditions were optimized using plasmids harvested from strain 5GB1C, and the detailed optimized procedure is described in Materials and Methods. Cells were harvested from both agar plates and liquid culture, but only cells harvested from agar plates yielded transformants. Osmotica are usually added into electroporation solutions to maintain osmotic balance and thus typically increase DNA transfer efficiency. Unexpectedly, all tested osmotica significantly reduced the transformation efficiency of strain 5GB1C. The transformation frequencies for Milli-Q H₂O, 9.3% sucrose, 10% glycerol, and 30% PEG 6000 were 5 × 10⁵, 10⁴, 5 × 10³, and 0 CFU per μg DNA, respectively (Fig. 1A). As shown in Fig. 1B, when 1-mm-gap cuvettes were used, field strengths of from 13 to 20 kV/cm gave similarly high electroporation transformation frequencies (∼5 × 10⁵ CFU per μg DNA). On the other hand, both a lower field strength (7 kV/cm) and a higher field strength (25 kV/cm) resulted in low electroporation transformation frequencies, possibly due to inefficient breakdown of the cell envelope or cell lysis, respectively. In addition, efficient transformation (>5 × 10⁵ CFU per μg DNA) was also obtained using 2-mm-gap cuvettes at a field strength of 12.5 kV/cm (data not shown). Following electric shock, methanol or methane was added as a carbon and energy source for cell recovery. Strain 5GB1C grows well in methane or methanol (0.2% to 2%, vol/vol), but as shown in Fig. 1C, methane was more favorable than methanol for cell recovery. The frequencies of positive colonies obtained when methane was used were at least 40% higher than those obtained when methanol was used. In addition, 0.2% methanol in the recovery medium led to obvious cell lysis and low transformation frequencies. We determined that the optimal methanol concentration for cell recovery is 0.02%. During optimization of the cell recovery time, we discovered that the highest frequency (∼5 × 10⁵ CFU per μg DNA) of transformants was obtained after overnight recovery (∼16 h), but the frequency of transformants obtained after 4 h of recovery (∼3.5 × 10⁵ CFU per μg DNA) was high enough for most genetic manipulations. Lastly, we investigated the storage of electrocompetent cells. Usually, electroporation-competent cells can be stored at −80°C and maintain a high electroporation efficiency for several months. However, frozen cells appeared to lyse after thawing, and it was not possible to identify conditions that would allow storage of electroporation-competent cells. The electroporation protocol utilized for all subsequent experiments with strain 5GB1C involved plate-grown cells, no osmoticum, 1-mm-gap cuvettes, settings of 25 μF and 200 Ω, and overnight growth on methane.

FIG 1.

DNA transformation frequencies under different electroporation conditions. The effects of the electroporation solution (A), the field strength (B), and the carbon source in the recovery medium and the recovery time (C) on DNA transformation frequencies in M. buryatense 5GB1C are shown. Plasmid pAWP89 harvested from strain 5GB1C was used in all experiments. Electroporation was performed using 1-mm-gap cuvettes and a Gene Pulser II system (Bio-Rad). Data represent the means from 3 replicates ± standard deviations. MeOH, methanol.

M. buryatense gene knockout via direct electroporation of a PCR product.

Direct electroporation of PCR-generated fragments containing an antibiotic resistance marker is a convenient gene transfer method for both chromosomal insertion and gene deletion applications. We showed that this approach could be used for efficient deletion of a control gene previously deleted through a conjugation-based approach (glgA1) (9), a gene regulating expression of the soluble methane monooxygenase, mmoD, and the whole sMMO operon (10 kb). The kanamycin resistance marker and two flanking homology regions were assembled by fusion PCR, purified by use of a PCR purification kit, and then directly electroporated into strain 5GB1C. The resulting mutants were verified by PCR amplification of the deleted region (Fig. 2A) and DNA sequencing (data not shown). The deletions of the mmoD gene and sMMO operon were also confirmed by a naphthalene plate assay (14) (Fig. 2B). The result for mmoD is in agreement with results indicating that MmoD is an essential activator of sMMO operon transcription in Methylosinus trichosporium OB3b (24). No difference in growth was observed for either the mmoD or the sMMO operon mutants (data not shown). The frequency of these homologous recombination events was negatively correlated with the length of the deleted region. When ∼800 bp of flanking DNA (on each side) was used, the deletion frequencies for mmoD (gene length, 0.37 kb), glgA1 (gene length, 0.83 kb), and the sMMO operon (operon length, 10 kb) were approximately 8.2 × 10³, 5.3 × 10³, and 3.7 × 10² CFU per μg DNA, respectively.

FIG 2.

Confirmation of the mutations delivered into strain 5GB1C. (A) Validation of the M. buryatense mutations generated in this study by PCR amplification. All fragments were further validated by sequencing. (B) Detection of the sMMO activity of strains 5GB1C, 5GB1C-ΔmmoD::km, and 5GB1C-Δsmmo::km. Cells grown in NMS2 medium without copper were subjected to a colorimetric plate assay developed by Graham et al. (16). The cells expressing sMMO turn deep purple when successively exposed to naphthalene and o-dianisidine. WT, wild type.

Electroporation of additional type I methanotrophs.

After our success with the optimization of an electroporation protocol with M. buryatense 5GB1C, we investigated whether this method could be applied to other type I methanotrophs. Two other type I methanotrophs of other genera, Methylomonas sp. LW13 and Methylobacter tundripaludum 21/22 (13), were tested by use of the protocol optimized for 5GB1C outlined above and in Materials and Methods, which omits osmotica and includes an overnight grow-out period. In contrast to M. buryatense 5GB1C, both of these strains grow at neutral pH and do not require NaCl for growth.

M. tundripaludum 21/22 cells grown in liquid culture were efficiently electroporated with both an E. coli-harvested replicable plasmid (pAWP78) and PCR-generated fragments to mutate glgA2 at 2.3 × 10³ CFU per μg DNA and 2.3 × 10⁴ CFU per μg DNA, respectively. Cells harvested from agar plates did not yield any transformants when they were electroporated with a plasmid harvested from E. coli but demonstrated a transformation efficiency of 8 × 10³ CFU per μg DNA when they were electroporated with the PCR fragment. Methylomonas sp. strain LW13 cells grown both in liquid culture and on agar plates were efficiently electroporated with PCR-generated glgA deletion fragments but not plasmid pAWP78 harvested from E. coli. The transformation efficiency of cells isolated from liquid culture was 3.7 × 10⁴ CFU per μg DNA; cells isolated from an agar plate demonstrated a transformation efficiency of 1.1 × 10⁵ CFU per μg DNA. The resulting glgA mutants were verified by PCR amplification (see Fig. S1 in the supplemental material) and DNA sequencing (data not shown). As in strain 5GB1C, attempts to transform frozen electrocompetent cells prepared from both plate and liquid cultures were unsuccessful with both Methylomonas sp. strain LW13 and Methylobacter tundripaludum 21/22. Taken together, our results show that this electroporation protocol can be applied to three phylogenetically and physiologically divergent type I methanotrophs. It seems likely that a similar approach of omitting osmotica and including a subsequent grow-out period will be successful in other type I methanotrophs, as long as variables such as liquid versus agar plate growth and plasmid versus PCR products are tested. These results suggest that the optimal DNA transfer frequency obtained in strain 5GB1C without osmotica was not due to the high pH optimum and the NaCl requirement of this strain, since DNA transfer was also successful in neutrophilic methanotrophs that grow with no added NaCl. The reason for the negative effect of the osmotica is not known but may involve a special cell structure or negative interactions of the osmotica with cellular metabolism during the recovery period.

sacB-based marker-free deletion strategy in M. buryatense 5GB1C.

As we are developing M. buryatense 5GB1C for use as an industrial strain, we focused on this methanotroph to identify additional genetic manipulation tools that can be used in conjunction with the electroporation method. The creation of unmarked deletions is more desirable than marker exchange for metabolically engineered strains, to allow subsequent genetic manipulations. To this end, a new positive and negative selection cassette (PNSC) was constructed. A zeocin resistance marker, designated Zeo^r (475 bp), was assembled by fusing P_tac, the SD sequence of lacZ, and the Sh ble gene together (18). The small size of Zeo^r facilitates the goal of a small construct for foreign DNA insertion. The fragment containing the native SD sequence and sacB gene was introduced 3′ of Zeo^r through fusion PCR, generating a short PNSC named the ZS cassette, which is only 1.9 kb (Fig. 3). The ZS cassette-based marker-free deletion strategy (Fig. 3) contains two key points. First, a 450-bp fragment just downstream of the target region to be deleted is used as the direct repeat sequence (DR) and is added ahead of the ZS cassette; second, the DR and ZS cassette are inserted just upstream of the target region without any deletion (the target region together with the ZS cassette is excised via recombination between DRs). This strategy was tested by deleting mmoD, glgA1, and the sMMO operon. Two flanking regions (∼800 bp on each side), DR, and the ZS cassette were fused by PCR and electroporated into strain 5GB1C. The frequency of the initial double-crossover event was roughly 600 CFU per μg DNA. Moreover, the second recombination event between the elongated DRs was efficient, with approximately half of the sucrose-resistant colonies undergoing target excision. This approach is thus more appropriate for the deletion of large regions of DNA and gives higher overall frequencies for small deletions as well.

FIG 3.

Scheme for sacB and PCR-based marker-free DNA fragment deletion in strain 5GB1C. To delete a target fragment (dotted line), a 450-bp region just downstream of target fragment is used as a direct repeat sequence (DR) and is put ahead of the Zeo^r/sacB (ZS) cassette. Transformants with an insertion of the fragment containing the DR and ZS cassette just upstream of the target region are selected on zeocin. Recombination between two DR sequences excises both the ZS cassette and the target fragment, and the resulting mutant is selected on 2.5% sucrose. P_tac, tac promoter; SD, Shine-Dalgarno sequence; Sh ble, zeocin resistance gene; LF, left flanking region; RF, right flanking region; Chr, chromosome.

Removal of the marker gene using the FLP-FRT system and introduction of a foreign fragment into the M. buryatense 5GB1C chromosome.

We developed a method parallel to sucrose counterselection for creating unmarked genetic manipulations that further decreases the size of the PCR products required to generate our desired construct. We chose to employ a site-specific recombinase system (Cre-Lox or FLP-FRT) for deleting the antibiotic resistance marker (25, 26). Surprisingly, despite the lack of a Cre recombinase homolog in the M. buryatense 5G genome, we found that LoxP sites were cleaved in the absence of an exogenous copy of Cre recombinase, implying that an enzyme in strain 5GB1C could catalyze recombination between two LoxP sites (data not shown). We turned instead to the FLP-FRT site-specific recombinase system for creating controlled, unmarked genetic manipulations.

As a proof of concept, we created a PCR insertion fragment containing the promoter of the pmo operon fused to xylE to be targeted into the 5GB1C chromosome between the METBUDRAFT_2794 and METBUDRAFT_2795 genes (Fig. 4). This region is transcriptionally silent (data not shown), allowing regulated transcription of heterologous genes inserted into the region. A fragment consisting of the Zeo^r marker flanked by two FRT sites was inserted into this construct. The resulting PCR product was electroporated into strain 5GB1C, and colonies with the correct insertion were isolated. To excise the Zeo^r marker by transiently expressing FLP, a replicable plasmid (pFC25) containing the flp gene under the control of P_tac was isolated from strain 5GB1C and electroporated into the insertion strain (the transformation efficiency of this plasmid is stated above). After the excision of the Zeo^r marker was confirmed (Fig. 2A), plasmid pFC25 was cured by passaging the modified strain in the absence of antibiotics to obtain the final strain. All originally tested zeocin-resistant transformants harboring pFC25 lost the Zeo^r marker, and all the zeocin-resistant transformants without pFC25 still contained the Zeo^r marker. Therefore, the 100% marker removal efficiency in strains bearing pFC25 indicates that the FLP-FRT system can be used in strain 5GB1C to efficiently unmark genetic manipulations in a controlled manner. In addition to introducing a target gene into the chromosome, the Zeo^r/FLP-FRT system can be applied to generate marker-free deletions.

FIG 4.

Scheme for FLP-FRT and PCR-based marker-free integration of a foreign fragment into the chromosome of strain 5GB1C. Four fragments of two flanking regions, the foreign DNA, and the zeocin resistance marker (Zeo^r) bordered by two FRT sites are assembled by PCR and inserted at the target site in the chromosome of strain 5GB1C using electroporation. Then, plasmid pFC25 containing the flp gene under the control of the tac promoter is electroporated into the insertion strain. After excision of the Zeo^r marker, pFC25 is cured by passaging the modified strain in the absence of antibiotics to obtain the final mutant. LF, left flanking region; RF, right flanking region; Chr, chromosome.

Deletion of R-M systems.

As mentioned above, 10 R-M system-associated restriction endonuclease genes (see Table S2 in the supplemental material) are predicted in the genome of M. buryatense 5G, and these may be a key factor limiting the transformation of foreign DNA into strain 5GB1C. To eliminate the obstacle of R-M systems, individual deletions in each of the 10 restriction endonuclease genes were attempted. Nine of the 10 deletions were successful. Only one (METBUDRAFT_1475) of these nine mutations led to the successful electroporation of a plasmid from E. coli, but it occurred at a low frequency (approximately 10 CFU per μg DNA). Interestingly, we could not delete the restriction endonuclease gene METBUDRAFT_0047, even after trying different flanking regions. Moreover, the attempt to delete this whole R-M system, including genes METBUDRAFT_0046 (methyltransferase gene) and METBUDRAFT_0047, failed as well. The methyltransferase of this R-M system was heterogeneously expressed in E. coli and purified (data not shown). However, an E. coli plasmid modified in vitro by this purified methyltransferase was still not successfully electroporated into strain 5GB1C. Because these nine genes were deleted separately, it is possible that more than one restriction endonuclease is a barrier to DNA transformation or that the one gene (METBUDRAFT_0047) that we could not delete is the main barrier. Therefore, in the future, a mutant lacking all restriction endonuclease genes except METBUDRAFT_0047 needs to be constructed to determine if the resulting strain will accept DNA from foreign sources.

Conclusions.

The bioconversion of natural gas/renewable biogas into valuable products is considered to be a promising strategy to utilize methane. To metabolically engineer methanotrophic strains for new traits, simple and rapid genetic tools are desired. A highly efficient DNA transfer method is the basis for developing rapid genetic tools. Therefore, we developed a robust electroporation procedure for three type I methanotrophic strains, a group for which electroporation has not previously proven successful. The protocols developed here streamline genetic manipulations in type I methanotrophs compared to previous methods of DNA transfer based on conjugation and enable the direct selection of double-crossover insertions and deletions, as well as the direct transfer of genes that might be toxic in other hosts. This is especially beneficial when engineering strains for industrial use, as multiple genetic manipulations often need to be combined in a single strain. Together with previous genetic tools that we have developed (9), a robust genetic manipulation system has been established for the fast-growing methanotroph M. buryatense 5GB1C, further enhancing its promise as an industrial organism.